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ABSTRACT
ADENOSINE TRIPHOSPHATASE ACTIVITY OF PLASTID MEMBRANE
DURING CHLOROPLAST DEVELOPMENT OF KIDNEY BEAN LEAVES
BY
Chen—hsiung Wu

2

l. The activity of a Ca +—dependent, trypsin—

activated ATPase from the membranes of kidney bean (Phaseolus

 

vulgaris) leaf plastids has been followed during greening.

2. In continuous light, there was no increase in
the total activity per leaf of this etioplast membrane
ATPase during the first 6 hours of plastid development. The
increase in ATPase activity occurred in the later stages of
chloroplast development.

3. The highest specific activity of the ATPase of
plastid membrane (based in umoles phOSphate released/mg
protein) was obtained from preparations of etioplast membrane.
In continuous light, specific activity of etioplast membrane
ATPase was constant for 9 hours, then decreased.

4. The relations between chlorophyll synthesis and
increase in plastid membrane protein and ATPase activity were
investigated under several different light regimes. Chloro-

phyll and plastid membrane protein could be increased without

Chen—hsiung Wu

concomittant increases in ATPase activity; but the converse
could not be obtained.

5. Pre—illumination of intact plants shortened the
lag phase of chlorophyll synthesis, and increased the amount
of plastid membrane protein and plastid membrane ATPase
activity immediately during subsequent continuous illumina—
tion.

6. Etiolated plants, treated with repetitive cycles
of 5 minutes of light and 8 hours of dark, showed an increase
in the total leaf ATPase activity after 5 cycles of light-
dark period while chlorophyll increased linearly with the
number of cycles and plastid membrane protein increased after
two light-dark cycles.

7. The ATPase of the etioplast membranes was more
easily inactivated by being kept at 55°C than the ATPase of
the chloroplast membrane.

8. ATPase was activated more by a combined treatment
of trypsin and dithiothreitol than by the treatment of tryp-
sin and dithiothreitol alone. The maximum activity of the
ATPase was obtained by a combined treatment of first dithio—
threitol, then trypsin; the reducing agent potentiates the
activation by trypsin but the reverse is not observed. This

indicates the mechanisms of ATPase activation by trypsin and

Chen-hsiung Wu
dithiothreitol are different.
9. I conclude that plastid membrane development and
increase in plastid membrane ATPase activity during chloro-
plast greening are not limited by chlorophyll synthesis.

The binding of this ATPase to plastid membranes appears to

change during greening.

ADENOSINE TRIPHOSPHATASE ACTIVITY OF PLASTID MEMBRANE
DURING CHLOROPLAST DEVELOPMENT OF KIDNEY BEAN LEAVES

BY

Chen-hsiung Wu

A THESIS

Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of

MASTER OF SCIENCE

Department of Botany and Plant Pathology

1974

[5‘ ACKNOWLEDGMENTS
\

I wish to thank Dr. Kenneth D. Nadler for the
inspiration and guidance he provided in the course of these
studies as my research supervisor. I would also like to
thank Dr. Clifford J. Pollard, Dr. Peter C. Wolk and
Dr. Loran L. Bieber for their service on my guidance
committee and Dr. Robert S. Bandurski for the use of his

facilities.

ii

TABLE OF CONTENTS

ACKNOWLEDGEMENTS
TABLE OF CONTENTS . . . . . . . . . . . . . . . . . . .
LIST OF TABLES . . . . . . . . . . . . . . . . . . . .
LIST OF FIGURES . . . . . . . . . . . . . . . . . . . .
INTRODUCTION . . . . . . . . . . . . . . . . . . . . .
MATERIALS AND METHODS . . . . . . . . .

Plant GrOwth . . . . . . . . . . . . . . . . . . . .
Isolation of Plastid Membrane . . . . . . . . . . . .
Assay of ATPase . . . . . . . . . . . . . . . . . . .

RESULTS 0 O O O O O O O O O O O O O O O O O O O O O O 0

ATPase of Plastid Membrane of Kidney Bean . . . . .
Trypsin Activation of the Plastid Membrane Ca2+—
dependent ATPase . . . . . . . . . . . . . . . . .
ATPase Activity of Plastid Membrane During
Chloroplast Development . . . . . . . . . . . . .
ATPase Activity During Chloroplast Development of the
Pre-illuminated Leaves (Which were Treated with 10
Minutes Light and Put in the Dark for 25 HCurs) .
Plastid Membrane ATPase Activity During Repetitive
Light— Dark Treatment . . . . . . . . . . . . . .
The Effect of Dithiothreitol and Trypsin on ATPase
Activity of Chloroplast Membrane . . . . . . . . .
Heat Inactivation of Plastid Membrane ATPase . . . .

DISCUSSION 0 O O O O O O O O O O O O O O O O O O O O .
LIST OF REFERENCES . . . . . . . . . . . . . . . . . .

APPme . O O O O O O O O O O O O O O O O O O O O O 0

iii

Page
ii
iii

iv

11

13
14

15
38

42

Table

LIST OF TABLES
Page

The effect of trypsin and dithiothreitol (DTT)
on ATPase activity of Chloroplast membrane . . . 36

Effect of pre-illumination on the Ca2+-dependent

ATPase activity and membrane protein of plastids
during chloroplast development . . . . . . . . . 37

iv

LIST OF FIGURES

Figure

1.

2.

10.

11.

The Ca2+-dependence of the ATPase activity of
chlorOplast membranes . . . . . . . . . . . . . .

The effect of the amount of etioplast membrane
protein on ATPase activity . . . . . . . . . . .

Trypsin activation of etioplast membrane ATPase .

Ca2+-dependent ATPase activity, chlorophyll
content and insoluble protein content of plastids
during chloroplast development in 12 days dark-
grown bean leaves . . . . . . . . . . . . . . . .

Long-term variations in Ca2+-dependent ATPase
activity, chlorophyll content and plastid membrane
protein in plastids greening bean leaf . . . . .

Effect of pre—illumination on the Ca2+-dependent
ATPase activity, chlorophyll content and insoluble
protein of plastids during chloroplast development

Effect of pre-illumination on the Ca2+—dependent
ATPase activity, chlorophyll content and insoluble
protein of plastids during chloroplast development

Conversion of protochlorophyllide a to chlorophyll
in etiolated leaves . . . . . . . . . . . . . . .

Regeneration of protochlorophyllide a after illu-
mination of etiolated leaves with 5 minutes light

Ca2+-dependent ATPase adtivity of plastid membrane
after several light-dark cycles of 5 minutes light
and 8 hours darkness . . . . . . . . . . . . . .

Time course of heat inactivation of trypsin-
activated Ca2+-dependent ATPase of etioplast and

chloroplast membranes . . . . . . . . . . . . . .

V

Page

23

24

26

28

29

30

31

32

33

35

INTRODUCTION

The formation of the lamellar system in plastids of
dark—grown leaves progresses through a series of light—
dependent steps (13, 14). The tubes in the prolamellar body
constitute a specific arrangement of the membrane as part of
the starting material for the development of the chloroplast
lamellar system. When etiolated leaves are illuminated with
white light, protochlorophyllide a is reduced photochemically
to chlorophyllide a, and the tubes in the crystalline pro-
lamellar bodies are transformed and dispersed into the
primary lamellar layers which are parallel to each other (13,
14,32)._ The acquisition of photophosphorylation activity
and photosynthetic competence all occur within the first few
hours of illumination of etiolated leaves (11. 25, 28). As
both chlorophyll a and b accumulate, extensive formation of
grana takes place. Grana formation is correlated with the
rapid phase of chlorophyll synthesis (11, 14). The detailed
molecular composition of the membranes of prolamellar body
is little known. Little is understood about the changes

occuring when the membranes of prolamellar body become

capable of photosynthesis, nor is the role of chlorOphyll
in this assembly process understood. I decided to study
these prdblems in chloroplast membrane formation using the

2+-—dependent ATPase activity of these membranes as a

Ca
marker.

It has been shown that the treatment of chloroplasts
or subchloroplast particles with trypsin activates a Ca2+-
dependent ATPase. Treatment with dilute EDTA solution
solubilizes this ATPase and releases a coupling factor which
markedly stimulates photophosphorylation in partially
resolved subchloroplast particles (3, 4, 5, 7, 8, 9, 18, 29,
30). Racker and his associates have shown that a Ca2+-
activated ATPase activity is associated with spinach chloro-
plast, and this ATPase can be washed from the membrane by
dilute EDTA solution. The Chloroplast membrane after
extraction by EDTA is incapable of carrying on photosynthetic
phosphorylation. The EDTA extract of chloroplast membranes
exhibits a trypsin- and dithiothreitol-activated, Ca2+—
dependent ATPase activity. ‘When this EDTA extract is added
(to the depleted chloroplast membrane, the photophosphory—
lation of chloroplast membrane can be restored (5, 23).

Ca2+-activated ATPase activity is also present in etioplast

membranes. This etioplast ATPase appears to have identical

properties to the chloroplast ATPase: for example, NaCl-EDTA
extracts of etioplasts can restore photosynthetic phOSphory—
lation activity to the depleted green membranes of chloro-
plasts (15, 19).

These studies were initiated to probe plastid
membrane development by relating the development of membrane—
bound ATPase activity to membrane protein and chlorophyll
content during greening.

The choice of suitable plant leaf tissue for these
studies is of some importance. Racker et. a1. (23, 30) used
spinach for studying properties of chloroplast ATPase:
Lockshin et. a1. (19) used maize for measuring the specific
activity of membrane bound ATPase during Chloroplast develop-
ment. waever it is inconvenient to use maize for greening
experiments because of the gradient in age of the leaf cells
in this plant. While Spinach leaves are more uniform, it is
difficult to grow large amounts of leaf tissue in the dark.
Therefore leaves of kidney beans, which green up synchronous-
ly and can be easily grown in the dark were selected as

‘ plant material in these experiments.

MATERIALS AND METHODS

Plant Growth

Kidney beans (Phaseolus vulgaris) were sterilized in
1%.Na0Cl for 15 minutes. After washing out the NaOCl with
water, the sterilized seeds were sown in 2.5 liters vermi-
culite in plastic flats (length 54 cm, width 26 cm, depth
6.5 cm) containing 2.5 liters Hntner's nutrient solution (in
appendix). After one week growth in darkness at ZS'C, plants
were watered with one liter deionized water. Leaves of the
etiolated plants were harvested generally at 11 days to 13
days. Before 11 days, the primary leaves were still enfolded
within the cotyledons and after 13 days or longer, the
etiolated leaves showed long lag phase of chlorophyll synthe-
sis or greened poorly. Greening plants were obtained by
putting the dark-grown plants in 430 ft—candles white light-
obtained from four 20 W incandesent bulbs and 2 cool white

20‘W fluorescent lamps.

Isolation of Plastid Membrane

To Obtain plastid membranes, etiolated leaves or

green leaves were ground in a solution of 0.01 M NaCl and
0.05 M.Tris-HC1 at pH 8.0 by mortar and pestle. Preliminary
experiments indicated that more ATPase activity (per mg
protein) was obtained by using a grinding medium without
sucrose than one with sucrose. This was continued in all
experiments described. After filtering the homogenate
through 4 layers of cheeseclOth and 2 layers of miracloth,
plastid membranes were Spun down by centrifugation at 3000 x
g for 15 minutes. The pellet of plastid membranes was resus—
pended in solution of 0.05 M Tricine-NaOH and 0.01 M NaCl at
pH 8.0 by the mixer. This suspension was used for assaying
Ca2+-dependent ATPase activity.

All experimental manipulations in the isolation of

intact plastids and plastid membrane were performed at ice

temperatures (0 - 4°C).

Assay of ATPase

 

For assay of ATPase activity, 0.5 ml plastid membrane
suspension was added to 2 m1 substrate containing 100 umoles
Tricine-NaOH at pH 8.0, 25 nmoles Ca2+ and 10 umoles ATP.

The reaction mixture was incubated for 30 minutes at 37°C in
a water bath and was then stopped by addition of 1 ml of

cold 16% trichloroacetic acid. The phOSphate released by

ATP hydrolysis during incubation was determined. After
Spinning out the TCA precipitate, the phosphate content
the supernatant was measured with the Ames reagent by
incubating at 45°C for 10 minutes and then reading the
absorbance at 820 nm with Gilford spectrophotometer 240
The non-enzymatically released phosphate was subtracted
all readings.

Protein content was determined by the method of
Lowry et. al. (20).

Chlorophyll content in the leaf was measured by

method of Arnon (2).

of

(1).

from

the

RESULTS

ATPase of Plastid Membrane of Kidney Bean
The Ca2+-dependence of the ATPase of etioplast or
chloroplast membranes of kidney bean leaves is shown in

2+

Fig. 1. An optimum Ca concentration occurs in the range

5 - 17.5 millimolar Ca2+. There was a slight inhibition at

25 millimolar Ca2+

in reaction mixture. The Specific
activity of the plastid membrane ATPase from 12 days old dark—
grown bean leaves as high as 9.4 umples ATP hydrolyzed per

mg membrane protein in 30 minutes could be obtained by

2+ and

incubating plastid membrane at 37'C with ATP, Ca
Tricine-NaOH buffer at pH 8.0. Levels of trypsin-treated
ATPase activity could be Obtained as high as 20.5 umoles per
mg membrane protein per 30 minutes at the same condition as
those without trypsin treated ATPase. This activity is low
compared with that from Spinach leaves as reported by Racker
et. a1. (23, 30) and higher than from corn leaves used by
Lockshin et. a1. (19). The Specific activity (on a mg

protein basis) of the ATPase of chloroplast membrane was

less than that of the etioplast.

7

Trypsin Activation of the Plastid Membrane
Ca2+-dependent ATPase

ATPase activity is linearly dependent on added
plastid membrane protein to at least 0.6 mg membrane protein
(Fig. 2). Trypsin activates this ATPase activity (Fig. 3).
A fifteen—minute incubation at room temperature with 10 Mg
trypsin yields the maximum ATPase activity. If 10 pg
trypsin are added 15 minutes after the first trypsin addition,
there is no additional increase in ATPase activity (Fig. 3).
In fact, the ATPase activity decreases slightly 10 minutes
after the second trypsin addition probably due to trypsin
digestion of some active ATPase. This indicates that 10 pg
trypsin treatment for 15 minutes at room temperature fully

activates the ATPase in 200 ug plastid membrane.

ATPase Activity of Plastid Membrane
During Chloroplast Development
The development of twelve-day old dark-grown etiolated
bean plants illuminated continuously with white light is
shown in Fig. 4. The lag phase of chlorophyll synthesis is
ended after 4 hours light. An increase in plastid membrane
protein per leaf could be detected only after 6 hours light.

Increases in total leaf ATPase activity, either with or

without trypsin activation, could also be detected only
after 6 hours light; however specific activity (expressed as
pmoles of phosphate released from ATP per mg membrane protein
in 30 minutes) decreased after 9 hours light for both trypsin
activated ATPase and non-trypsin ATPase.

After 3 days of continuous light, the total ATPase
of chloroplast membrane and Chlorophyll in the leaf remained
constant, but membrane protein of chloroplast still increased
for 3 more days in continuous light (Fig. 5). The data of
Fig. 5 also Show that ATPase of plastid membranes, the
plastid membrane protein and chlorOphyll all increased

maximally between 1 day and 3 days continuous light.

ATPase Activity During,Chloroplast Development of
the Pre-illuminated Leaves (Which were Treated with 10
Minutes Light and Put in the Dark for 25 Hours)

The lag phase of chlorophyll synthesis can be
shortened or abolished by treating the plant with a short
light period followed by a long period of incubation in
darkness (31, 33). This long incubation in darkness after
brief illumination probably permits the enzymatic adaptation
of the etioplast; i. e. the brief illumination followed by

a period of prolonged darkness results in the synthesis of

10

some of the enzymes necessary for ALA formation (24) and
photosynthesis (21). I therefore studied the effect of pre—
illumination on plastid membrane ATPase activity during
greening.

The etiolated leaves of 12 days dark-grown plants
were illuminated for 10 minutes and put in the dark for 25
hours, then were illuminated with continuous light. Compared
to non-preilluminated experiment shown in Fig. 4, this
treatment markedly shortens the lag phase of chlorophyll
synthesis and results in slight but immediate increases in
the membrane protein of plastid per leaf and total ATPase,
both trypsin-activated and non-trypsin-activated, activity
of plastid membrane per leaf. The specific activity of the
ATPase decreases immediately after exposing pre-illuminated
plants to continuous light (Fig. 6 and Fig. 7) as opposed to
the relatively constant ATPase activity of control plants,
not pre-illuminated.

I have repeated this pre-illuminated treatment
experiment. The data for plastid membrane protein per leaf,
total ATPase activity of plastid membrane per leaf and
Specific ATPase activity of plastid membrane are represented
in Table 2. There is no doubt that plastid membrane protein

content per leaf increases and the specific activity of

11

plastid membrane ATPase immediately decreases after the
beginning of the continuous light. Although there is no
significant increase in total ATPase activity per leaf during
the first two hours of illumination, the total ATPase content
of the leaf linearly increases with illumination time beyond
2 hours. I believe that there is an immediate slight
increase in total ATPase activity per leaf after the begin—

ning of the continuous light after pre—illumination.

Plastid Membrane ATPase Activity During
Repetitive Light-Dark Treatment
If etiolated leaves are repetitively illuminated
with 5 minutes light, then returned to darkness for several
hours, this treatment can induce the etioplast of the dark—
grown leaves partially develop into a chlorOplast, although
chloroplasts developed under this condition lack grana
formation (6, 10, ll, l3, 14, 28). Such chloroplasts show
large stacks of the parallel primary thylakoids with very
little fusion. These plastids have as much protein as mature
chloroplasts. In order to select conditions for the light
part of the treatment, I determined how much time was
required to convert all proto-chlorophyllide a in etiolated

leaves to chlorophyllide. In the 12 days dark-grown

12

etiolated bean leaves, this required 3 minutes illumination
(Fig. 8). In fact, most of protochlorOphyllide a was photo—
converted to chlorOphyll within 1 minute. After 1 minute,
there is only a small amount protochlorophyllide a in the
etiolated leaves but this small amount of protochlorophyllide
a requires an additional another 2 minutes illumination for
complete conversion to chlorOphyllide.

In order to ensure that all protochlorophyllide a in
the etiolated leaves had been converted to chlorophyllide by
brief light, five minutes instead of three minutes illumina-
tion of the etiolated plants was used. I then determined
the length of the dark period needed for resynthesis of the
maximum amount of protochlorophyllide a in the etiolated
leaves. This required 7 hours darkness (Fig. 9). In the
first two hours darkness after the brief illumination, there
is little protochlorophyllide a Synthesis, as in leaves with
a lag phase in Chlorophyll synthesis. Eight hours were used
for the dark period in this experiment for reasons similar
to those mentioned above. Thus, the dark—grown leaves of
kidney bean were treated repetitively with cycles of 5
minutes light and 8 hours darkness.

The results of this treatment are indicated in Fig.

10. Leaf chlorophyll content increased linearly for 6

13

light-dark cycles. This means that the rate of chlorophyll
synthesis and therefore protochlorophyllide regeneration
within 6 light-dark cycles remains constant. After 6 cycles
of light-dark period, the rate of chlorOphyll synthesis
increases greatly indicating that the rate of ALA synthesis
is now greater than that before 6 light—dark cycles.

Plastid membrane protein and leaf fresh weight increase after
2 cycles of light-dark period while plastid membrane ATPase

increases only after 5 cycles of light-dark period.

The Effect of Dithiothreitol and Trypsin on
ATPase Activity of Chloroplast Membrane

The plastid membrane ATPase activity can be activated
by several methods; by treatment with trypsin, by heating two
minutes at 65°C in the presence of ATP and by sulfhydryl
compounds such as dithiothreitol, a-thioglycerol and p—
mercaptoethanol (23, 30). The ATPase activity of chloro—
plast membrane treated by dithiothreitol for 2 hours was
much less than those activated by trypsin (Table 1). If
ATPase was activated by trypsin for 15 minutes, then acti-
vated by dithiothreitol for one hour or two hours, or if
the ATPase was activated by dithiothreitol for one hour or

two hours, then activated by trypsin for 15 minutes, the

14

resulting ATPase activity was more than that obtained by
trypsin— or dithiothreitol-activation alone. Furthermore,
the activity of ATPase first activated by trypsin then by
dithiothreitol was about equal to the combined activities of
ATPase activated by trypsin and dithiothreitol alone.
Hewever the activity of ATPase first activated by dithio—
threitol then by trypsin was much more than the activity of
ATPase first activated by trypsin then by dithiothreitol.
Clearly there is a synergistic effect of dithiothreitol on

trypsin activation.

Heat Inactivation of Plastid Membrane ATPase

 

A change in the thermal stability of a protein indi-
cates a change in protein conformation and/or a change in the
environment of a protein. The ATPase of etioplast membranes
is rapidly inactivated by 55°C (Fig. 11), while the ATPase of
chloroplast membranes is relatively stable at 55°C. Although
the ATPase of chlorOplast membrane drops to approximately 80%
of the control ATPase after 10 minutes incubation, the ATPase
activity of etioplast membrane drops to 30% of the control
ATPase after 10 minutes incubation. The different heat sensi—
tivity between etioplast membrane ATPase and chloroplast
membrane ATPase may indicate that the environment of plastid

membrane ATPase changes during chloroplast development.

DISCUSSION

ATPase of crude bean plastid membrane preparation is
used for assaying the ATPase activity in these experiments.
Mitochondria also have trypsin activated, Ca2+-dependent
ATPase (30). Using an isolation medium without sucrose to
isolate plastid membranes and using a low Speed centrifugation,
contamination by mitochondrial membranes was minimized.

Lockshin et. al. measured plastid membrane ATPase
activity of maize leaves during greening. ATPase activity
per mg membrane protein was relatively constant through 12
hours of illumination. After 12 hours of illumination, the
specific activity of ATPase decreased by a factor of about 2
during the next 13.5 hours regardless of whether or not the
plants were illuminated. They suggested that most or possibly
all of the coupling factor (measured as ATPase activity) of
the chlorOplast may be present in the etioplast from which
it develops. They also reported that the specific activity
of the enzyme dropped two fold during the next 13.5 hours due
to synthesis of much more membrane but little new synthesis

of the coupling factor occurs (19). This does not appear to

15

16

be the case in greening bean leaves. The specific activity
of the Ca2+-dependent ATPase of bean plastid membranes
remains constant in early period of greening because there

is neither net membrane protein synthesis nor ATPase synthe—
sis at this time: the specific activity of the ATPase dropped
in the later period of chloroplast greening since other
membrane proteins of plastids were synthesized much more than
the ATPase during this period. The total plastid membrane
ATPase of the leaf increases in continuous light, after a

lag period, for about 3 days. Horak and Hill (16) also
reported that there is a considerable increase in the Speci-
fic activity of the ATPase in plastids during greening. They
measured the activity of the ATPase in EDTA extracts of
plastids and found that it increased during greening. The
proteins extracted by EDTA from plastid membrane are mainly
ATPase with little contamination by other membrane proteins
(17).

All the ATPase activity of plastid membranes during
the first 6 hours in continuous light or in 5 light-dark
cycles is present in the etioplast. If the ATPase of the
etioplast membrane is originally in the prolamellar body (19),
it is undoubtedly redistributed into lamellae during the ‘

first several hours of chloroplast development.

17

The lag phase of chlorophyll synthesis ends within 4
hours during continuous illumination: ATPase activity of
plastid membranes increases only after 6 hours in continuous
light. Furthermore, in conditions where etiolated leaves
were treated by cycles of 5 minutes of light and 8 hours of
the dark, chlorophyll accumulates while the ATPase activity
of plastid membrane increases only after 5 cycles of light-
dark period. The above experimental results suggest that
(a) chlorophyll does not limit the synthesis of plastid
membrane ATPase, (b) ATPase synthesis can occur in the dark
without Simultaneous chlorOphyll synthesis (although chloro-
phyll synthesis must precede it).

There was no chlorOphyll formation (except that a
small amount of protochlorOphyllide a was converted to chlo-
rophyll a within the first 3 minutes of light) in etiolated
leaves during the 3 hours lag phase of chlorophyll synthesis.
Mego and Jagendorf (22) showed that the etiolated bean
plastids had the same amount of chlorophyll formed in one
minute or two hours of light: but plastid growth was greater
in two hours light than one minute light (22). By the use
of red light and far—red light to control enlargement of
bean leaf plastids, they also provided evidence that there

was no correlation between plastid size and the amount of

18

Chlorophyll development. This suggests that chlorOplast
membrane development is not controlled by chlorophyll synthe—
sis.

It has been shown by Butler and associates that when

the dark-grown bean leaves (Phaseolus vulgaris) were illumi—

 

nated with white light, photophOSphorylation appeared after
one to two hours of illumination (11, 28). Using similar
plants in these experiments, I have also shown that there is
no increase in chlorophyll and plastid membrane protein
content as well as ATPase activity in this period. Because
the membranes of prolamellar body progress through a series
of changes and arrangements that are stimulated by light, I
suggest that neither Chlorophyll nor plastid membrane protein
synthesis but some kind of conformational changes in prola-
mellar body membrane is required for photophOSphorylation
development.

Glydenholm and Whatley (12) also reported that '
cyclic and non-cyclic photophosphorylation were detectable
only after 10 hours illumination. In neither report did the
development of photophOSphorylation parallel the increase of
ATPase activity of etioplast membranes during chloroplast
development as was shown in my experiment. Even if the

ATPase of plastid membranes is related to a coupling factor

19

of photophOSphorylation, the time for the onset of photophos—
phorylation of etiolated leaves would not correspond to the
time for increase ATPase activity. Because there is already
a certain amount of ATPase in etioplast membrane, it can be
used for the development of photophoSphorylation without new
ATPase synthesis. Indeed, ATPase Synthesis may not commence
until the activity of etioplast ATPase becomes limiting for
photophosphorylation at later greening stages. So we can not
compare the time relationship between photophosphorylation
development and the ATPase synthesis to conclude whether the
ATPase of plastid membrane is a coupling factor or not.

The Ca2+adependent ATPase of plastid membrane is a
very stable enzyme. For the assay of ATPase activity, the
enzyme was incubated at 37°C for 30 minutes. In fact this
ATPase, in the presence of ATP, can be heated at temperature
as high as 65°C for 2 minutes to increase its activity (23,
30). This high stability of the ATPase of chloroplast
membranes may be due to the high thermal stability of chloro-
plast lamellae. The lamellar proteins and lipids may
protect it from denaturation by the environment (5). That
the ATPase of etiOplast membranes is easier to denature by
heat than the ATPase of chloroplast membrane suggests that

the synthesis or reorganization of the proteins and lipids

20

of chloroplast membranes occurs during greening.

Racker et. a1. first isolated a coupling factor
from chloroplasts. This coupling factor has been shown to
stimulate photophosphorylation and to have trypsin-activated,
Ca2+-dependent ATPase. They called it chloroplast coupling
factor 1 (the abbreviation used is CFl) to distinguish it from
mitochondrial coupling factor 1 (the abbreviation used is Fl)
(23, 30). Recently, Racker et. al. (26, 27) reported that
five different polypeptides could be isolated from CFl.
These five subunits are referred to as a—, 3—, 7—, 5- and e—
chains in the order of decreasing molecular weight of 59000,
56000, 37000, 17500 and 13000. They called e subunit a CFl
inhibitor; because it can inhibit ATPase activity. Antibodies
against the 59000 and 37000 molecular weight subunit (a and 7
component) inhibited cyclic photophosphorylation and phos-
phorylation coupled to the Hill reaction. They concluded
that a and 7 subunits of CFl are intimately involved in the
function of the coupling factor and the 6 subunit is a
regulatory subunit of the enzyme. They proposed that CFl
inhibitor is more firmly bound to CFl at a site required for
binding to the membrane. They suggested that trypsin activa—
ted the ATPase of CFl either by destroying the inhibitor or

removing it from the active site of the enzyme and dithio—

21

threitol somehow interferes with the interaction of the
inhibitor with the active site of CFl.

In my experiments, plastid membrane ATPase after
activation by trypsin can still be activated by dithiothrei-
tol. So, I don't support the suggestion by Racker et. al.
that dithiothreitol interferes with the interaction of inhibi—
tor with active site of CFl. Instead the evidence reported
here suggest that there are two reactions to activate the
enzyme ATPase. One is to reduce disulfide groups(-S-S—) of
protein to sulfhydryl groups (—ZSH). Another activation is
to remove CFl inhibitor. Thus the enzyme ATPase can enhance
its activity either when its disulfide groups are reduced to
sulfhydryl groups or when CFl inhibitor is removed. Thus
the enzyme ATPase after trypsin activation can still be
activated by dithiothreitol and vice versa. The reason for
the enhanced activity of ATPase first activated by dithio-
threitol then by trypsin is not clear. Perhaps the protein
conformation of the enzyme is affected by this treatment.
Racker et. al. suggest that trypsin activated Caz+-
dependent ATPase of chloroplast was associated with a solubi-
lization of the ATPase (23). If plastid membrane ATPase is
first treated by dithiothreitol, many disulfide bonds (—S-S—)

are reduced by dithiothreitol to sulfhydryl groups (—2SH).

22

This reaction many render plastid membrane protein more open
to removal of CFl inhibitor by trypsin; and the maximum
activity of ATPase may be obtained.

Racker and coworkers (23) have Shown that EDTA
extracts of chloroplast (CFl) could stimulate photophosphory-
lation. But when this extract was treated by trypsin to
exhibit ATPase activity, its properties as a coupling factor
were lost. Horak and Hill demonstrated that no coupling
factor activity could be detected in disc gel electrophoresis
zones containing ATPase activity (16). The trypsin-activated
ATPase can not function as a coupling factor. The ability
to couple requires CFl inhibitor binding to the ATPase.

Both ATPase activity and photophosphorylation could be easily
demonstrated in the chlorOplasts that have been exposed to
dithiothreitol (23). So, dithiothreitol-activated ATPase

contains the trypsin-activated ATPase and the CFl inhibitor.

23

 

 

 

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millimolar (Ca2+) in reaction mixture

Figure 1. The Ca2+-dependence of the ATPase activity of
chlorOplast membranes. 190 pg membrane protein of chloro—
plast were treated by 10 Mg trypsin for 15 minutes and
trypsin activation was stepped by adding 100 ug trypsin
inhibitor. Enzymatic phosphate release was measured as
described in Methods except calcium salts were not included
in the substrate stock solution.

24

 

ATPase activity (uMPi released/30 min.)
A)

 

J l I l I
0 0.2 0.4 0.6 0.0 1.0

Plastid membrane protein added (mg)

Figure 2. The effect of the amount of etioplast membrane
protein on ATPase activity. 30 pairs of primary leaves of
14 days dark~grown plants were ground in 20 ml of 0.05 M
Tris-HCl buffer containing 0.01 M.NaCl at pH 8.0 with sand
in a mortar and pestle. After filtering through 4 layers

of cheesecloth and 2 layers of miracloth, plastid membranes
were spun down by centrifugation at 3000 x g for 15 minutes.
The plastid pellet was suspended in 10 ml of 0.05 M Tricine—
NaOH buffer containing 0.01 M NaCl at pH 8.0 by the mixer.
Various amount of plastid membrane suspension were assayed
for ATPase as described in Methods.

25

d
0

UI

 

Spec1f1c actIVIty of ATPase
(uMPi released/30 min./mg protein)

 

I I I
o 10 20 so

Incubation time (min.)

Figure 3. Trypsin activation of etioplast membrane ATPase.
200 pg plastid membrane was activated by 10 ug trypsin and

10 ug trypsin was further added after 15 minutes incubation
as indicated by an arrow.

26

Figure 4. Ca2+-dependent ATPase activity, chlorophyll
content and insoluble protein content of plastids during
Chloroplast development in 12 days dark-grown bean leaves.
Plants were illuminated under continuous light. 10 pairs

of primary leaf were harvested each time and the plastid
membranes were isolated as described in the Methods. 0.05
ml membrane suSpension were activated by treatment with

10 pg trypsin for 15 minutes. Trypsin activation was

stopped by adding 100 ug trypsin inhibitor. Then, plastid
membrane suspension was assayed for ATPase as in the Methods.
(0) chlorophyll content. (A) total ATPase activity of
non-trypsin treated plastid membrane per leaf. (I) total
ATPase activity of trypsin treated plastid membrane per leaf.
(I!) specific activity of non—trypsin activated ATPase of
plastid membrane. (0) specific activity of trypsin activa-
ted ATPase of plastid membrane. (Cl) total plastid membrane
protein per leaf.

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Illumination time (hours)

28

 

 

 

 

Total activity of ATPase

 

 

 

 

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Illumination time (days)

Figure 5. Long-term variations in Ca2+-dependent ATPase
activity, chlorophyll content and plastid membrane protein

in plastids greening bean leaf. Methods for the experiment are
the same as in Fig. 4. (I) total ATPase activity of trypsin
treated plastid membrane per leaf. (A) total ATPase
activity of non-trypsin treated plastid membrane per leaf.

(0) total protein of plastid membrane per leaf. (0)
chlorophyll content.

Chlorophyll content (mg per leaf)

Specific activity of ATPase
(”MPi released/30 min./mg protein)

29

 

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N
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Total act1v1ty of ATPase(uMP1 released/30 min.)
GI
r

 

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Plastid membrane protein (mg per leaf)
L

 

 

o 2 4 6 8

Illumination time (hours)

Figure 6. Effect of pre—illumination on the Ca2+-dependent
ATPase activity, chlorophyll content and insoluble protein
of plastids during chloroplast development. Leaves were
illuminated for 10 minutes and placed in darkness for 25
hours, then continuously illuminated for various periods.
Methods for the experiment are the same as in Fig. 4.

(O) chlorophyll content. (I) total ATPase activity of
trypsin treated plastid membrane per leaf. (0) Specific
activity of trypsin activated ATPase of plastid membrane.
(CI) total protein of plastid membrane per leaf.

Chlorophyll content (mg/g fresh weight of leaf)

Specific activity of ATPase
(UMPi released/30 min./mg protein)

30

 

8
I
i released/30 min.)

B
I

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Total actIVIty of ATPase
‘1 .

 

I I I

Plastid membrape protein (mg per Leaf)
Chlorophyll content (mg/g fresh weight of leaf)

 

 

o 2 4 6 3

Illumination time (hours)
Figure 7. Effect of pre-illumination on the Ca2+-dependent
ATPase activity, chlorophyll content and insoluble protein

of plastids during Chloroplast development. Leaves were
illuminated for 10 minutes and placed in darkness for 25
hours, then continuously illuminated for various periods.
Methods for the experiment are the same as in Fig. 4.

(o) chlorophyll content. (A) total ATPase activity of
non-trypsin treated plastid membrane per leaf. (4A) specific
activity of non—trypsin activated ATPase of plastid membrane.
(Cl) total protein of plastid membrane per leaf.

31

40r-

Chlorophyll content
(pg/10 pairs of primary leaf)

 

 

01234567

Illumination time (min.)

Figure 8. Conversion of protochlorophyllide a to Chlorophyll
in etiolated leaves. 12 days old etiolated leaves were
illuminated and the pigments extracted as described previously.

32

 

 

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Time in the dark (hours)

Figure 9. Regeneration of protochlorophyllide a after
illumination of etiolated leaves with 5 minutes light.
Leaves were put in the dark after this 5 minutes light.
leaves were given for 5 minutes light again after every
hour in the dark and measurement of the increase in chloro-
phyll content used as an index of protochlorophyllide a
content.

33

Figure 10. Ca2+-dependent ATPase activity of plastid membrane
after several light-dark cycles of 5 minutes light and 8 hours
darkness. After each illumination, the leaves were assayed

as previously described. (0) chlorophyll content.

(4) total ATPase activity of non-trypsin treated plastid
membrane per leaf. (I) total ATPase activity of trypsin
treated membrane per leaf. (CI) total protein of plastid
membrane per leaf. (x) fresh weight of leaves.

34

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Figure 11. Time course of heat inactivation of trypsin—
activated Ca2+—dependent ATPase of etioplast and chloro—
plast membranes. The plastid suSpension in the reaction
mixture after heating was rapidly cooled in the cold ice
water. Total ATPase activity of etioplast membrane and
chlorOplast membrane before heating are 2.4 pmoles and 1.8
pmoles phosphate released per 30 minutes reSpectively.

(I) ATPase of chloroplast membrane. (0) ATPase of
etioplast membrane.

 

 

36

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LIST OF REFERENCES

LIST OF REFERENCES

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A PPEND IX

APPENDIX

For 2 liter Hutner's (-Mg2+) stock solution:

Solution A: in 400 ml distilled water

 

Ca(NO3)2 35.40 g
EDTA(acid) 50.00 g
KZHPO4 40.00 g
KOH(85% pellets) 24 - 26 9

Cool with ice.

Solution B: in 300 ml distilled water

ZnSO4-7H20 6.59 9
H3803 1.42 g
Na2M04-2H20 2.52 9
CuSO4-5H20 0.394 9

Add 1 N HCl until cloudiness disappears.
Solution C: in 100 ml
FeSO4-7H20 2.47 9
Then solution A + solution B + solution C + H20—>2 liters.
"Mg2+" stock solution: 25 g MgSO4-7H20 per liter
The complete nutrient solution for plant growth:

2+"

(1 part Hutner's solution + 1 part "Mg stock solution)

—>di1uted to 50 times by H20
42

HIGAN STATE UNIVERSITY LIBRAR

1293

in 3|

428

HES